Conventional diagnostic procedures for identifying viruses include seeding containers with particular cell lines selected on their sensitivity to certain viruses and then inoculating the cell culture with a biological sample putatively containing a virus. Such biological samples include among other things saliva, urine, feces, cerebrospinal fluid (CSF), respiratory fluids and swabs such as those from the mouth, nasal cavity, throat, skin and genitals. The inoculated cell culture is then incubated and the cells examined for cytopathic effects induced by the virus. As certain viruses only grow on certain cells, the virus can be identified on the basis of the cell type in which it either induces a cytopathic effect (CPE) or does not induce a cytopathic effect.
There are a number of alternative protocols to this procedure including subjecting cells which have been inoculated with a virus preparation by trypsonisation to remove the cells, followed by virus detection using monoclonal antibodies specific for viral-derived polypeptides which are labeled with a reporter molecule such as a fluorescein (FITC) molecule. A further alternative is to include a cover slip within a culture tube in order to enhance recovery of the cells.
The conventional (or traditional—drum) method utilizes screw cap tubes which are seeded with appropriate cell lines. After the cells reach about 80% confluency the tube is inoculated with an appropriate specimen and monitored for CPE for up to three weeks. Daily monitoring of CPE is required for the first week. Less frequent monitoring is necessary for the second and third weeks. Often, blind passage is required to enhance virus recovery.
One of the disadvantages of this method is that it is time and labour intensive because daily monitoring of the tubes is required. Generally, two people inspect the same tube for CPE by light microscope to avoid subjectivity. In addition, not all viruses cause a visible CPE and those which do not are unable to be detected by this method. Furthermore, CPE formation monitored in the conventional tube method is highly dependent on the sensitivity of the cell lines and the capability of the virus to produce visible CPE. Toxicity of the specimen may also disadvantageously produce changes similar to viral CPE giving a false result. Also, some viruses produce CPE only after a long period of time (for example, Cytomagalovirus (CMV)). Thus, as results obtained by the conventional tube method are predominantly based on CPE detection and are not routinely confirmed by any other method, inaccurate diagnosis can occur. Another limitation of the conventional tube method is that it is difficult to use more than 2 or 3 tubes per specimen due to the resulting accumulation of tubes. For example, 40 specimens per day would create 500 tubes to analyze in the first week alone.
The shell vial method is currently the most advanced method utilized by those in the art for virus recovery. This method employs the use of a 5 ml plastic vial (shell vial), 16 mm in diameter which has a translucent lid. Following an appropriate treatment, a round (13 mm) cover slip is inserted into the vial. The vial is then seeded with a sensitive cell line which grows a monolayer on the cover slip. When the cell monolayer reaches about 80-90% confluency, the medium is discarded, the monolayer inoculated with the patient's specimen and the vial incubated. Then, the incubated vial is monitored for CPE, followed by the removal of the cover slip. The slip can then be fixed to a microscope slide and stained with monoclonal antibodies.
The advantage of the shell vial method is that virus recovery can be enhanced by centrifugation of vials after inoculation which can shorten the length of time taken to obtain results to as little as 2-3 days. Further, using the shell vial method there is no need to wait for visible CPE. The cover slip can be removed on the second or third day and stained with appropriate monoclonal antibodies and results confirmed using antibody-antigen staining.
However, the shell vial method also has a number of disadvantages. It is time consuming as the cover slips require special treatment; such as multiple washings with detergent and acetone followed by washing in distilled water and sterilization. The cover slips also have to be manually inserted into the vials. Further, if immunofluorescent staining is necessary, the procedure becomes even more complicated and time consuming. The medium from the shell vial has to be discarded and the cover slip manually removed using specific forceps, air-dried and fixed to a microscope slide, using vacuum grease. The removal of cover slips is tedious, since the cover slips may be broken by rough manipulation or unintentionally turned and fixed to the microscope slide with the monolayer upside down. Another complication may arise if the seeded cells also grow on the bottom of the cover slip, thus causing the cover slip to fix to the vial and making removal of the cover slip very difficult. Practically, as for the conventional tube method, using the shell vial method it is impossible to use more than 2 or 3 tubes per specimen due to the accumulation of tubes (i.e. 40 specimens per day creates 500 shell vials per week). Further, a large amount of monoclonal antibodies is required for immunofluorescence staining in order to cover the round 13 mm cover slip.
The 96 well plate method is another method which is used only in limited cases for recovery of viruses which grow on the same cell line. For example, if the wells are seeded with LLC-MK2 cell line the recovery of parainfluenza and also influenza viruses is possible. The 96 well plate method has advantages in that it is relatively easy to inoculate seeded cell lines with a particular specimen. Further, a large number of specimens can be inoculated onto the same plate and enhancement by centrifugation is also possible. Still further, the 96 well plate method only utilizes a small amount of media (0.3 ml instead of 1-1.5 ml used in the shell vial method), antigen-antibody techniques may be used for confirmation of results and the method also enables easy to “read” monitoring of CPE.
However, the 96 well plate method also has its disadvantages. The whole plate must be used for antigen-antibody detection which is not generally practical and the entire plate has to be used on the same day, even when the number of specimens is smaller than required for the whole plate. This means that for each day a new set of different plates must be used. This disadvantageously results in a situation where, once the detection is completed, there are no remaining cells available for a repeat procedure in case of an error or after a prolonged incubation period. Further, commonly only one or two different cell lines can be used per plate and the same type of specimen inoculated onto the plate.
Single well methodology, as described in Australian Innovation Patent No. 2001100242, alleviates problems associated with the conventional methods described above and provides an alternative, effective and economical process for conducting viral diagnosis. The disclosure of that patent is incorporated herein in its entirety by reference.
Flat-based wells are known to provide certain advantages when used in the context of diagnostic assays. In particular, they provide for more precise analysis compared with, say, round or curved-based wells. However, simple flat-based wells also have a disadvantage in that a meniscus tends to form in the base of the well resulting in uneven distribution of solution across the base of the well. In light of this, the inventor has developed a flat-based well that alleviates this problem as described below.